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Summary

We developed a method to efficiently ablate a single cell type, the
zebrafish melanocyte, and study the mechanisms of its regeneration. We found
that a small molecule, (2-morpholinobutyl)-4-thiophenol (MoTP), specifically
ablates zebrafish larval melanocytes or melanoblasts, and that this
melanocytotoxicity is dependent on tyrosinase activity, which presumably
converts MoTP to cytotoxic quinone species. Following melanocyte ablation by
MoTP treatment, we demonstrate by BrdU incorporation experiments that
regenerated melanocytes are derived from the division of otherwise quiescent
melanocyte precursors or stem cells. We further show that larval melanocyte
regeneration requires the kit receptor tyrosine kinase. Our results
suggest that a small number of melanocyte precursors or stem cells unevenly
distributed in larvae are drawn upon to reconstitute the larval melanocyte
population following melanocyte ablation by MoTP.

INTRODUCTION

One of the enduring challenges of modern biology is a mechanistic
understanding of regeneration. In understanding the many strategies used by
organisms for regeneration, two important questions are the role of cell
division or growth, and the origin of the cells used to replace the missing
tissues. Historically, regeneration studies in systems such as the salamander
limb or hydra have revealed two major modes of regeneration, epimorphosis and
morphallaxis. The salient mechanistic distinction between them is the role of
cell division; epimorphosis involves active cellular proliferation and
morphallaxis is achieved through re-patterning of the remaining cells without
a role for growth or cell proliferation
(Morgan, 1901) (reviewed by
Lenhoff and Lenhoff, 1991).
More recently, attention has focused on the origins of cells contributing to
regeneration, revealing roles for both differentiated and undifferentiated
precursors in various modes of regeneration. This distinction, combined with
Morgan's original distinction based on growth and cell proliferation, suggests
four classifications for regeneration mechanisms subject to cell division and
differentiated state of regeneration precursors, as follows. (1) Regeneration
via direct cell division from differentiated cells
(Fig. 1A). This mechanism is
sometimes called compensatory regeneration, exemplified by mammalian liver
regeneration (reviewed by Michalopoulos
and DeFrances, 1997). (2) Regeneration via cell division from
undifferentiated precursors or stem cells
(Fig. 1B). This mechanism
closely matches the current usage of epimorphosis, and is best exemplified by
the stage of limb regeneration following formation of the blastema
(Chalkley, 1954;
Hay and Fischman, 1961). (3)
Regeneration without cell division by transdifferentiation from otherwise
differentiated cells, as occurs in hydra following resection
(Park et al., 1970;
Holstein et al., 1991)
(Fig. 1C). (4) A fourth, as yet
hypothetical, mode of regeneration is formally suggested by this
classification scheme and involves regeneration without cell division from
late-stage undifferentiated precursors
(Fig. 1D). We note that
additional mechanisms, such as dedifferentiation of differentiated cells to
form a blastema are also employed in some regenerating systems. Additionally,
regeneration schemes may combine some of these mechanisms, either sequentially
in the same lineage [for instance, in limb regeneration, dedifferentiation of
myofibers to form the blastema is then followed by proliferation and new
growth (Kintner and Brockes,
1984)] or use different mechanisms for the different cell types or
lineages contributing to regeneration of more complex tissues [for instance,
in zebrafish fin regeneration, keratinocytes divide directly
(Poleo et al., 2001), whereas
melanocytes are thought to be derived from undifferentiated precursors or stem
cells (Rawls and Johnson,
2000)].

Another opportunity for understanding regeneration and the underlying
mechanisms of regulation comes from the analysis of homeostasis of single cell
types in mammals, such as erythrocytes (reviewed by
Morrison et al., 1995),
epidermis (Jones and Watt,
1993) (reviewed by Fuchs and
Raghavan, 2002), and epithelium in the lining of the small
intestine (Cheng and Leblond,
1974) (reviewed Potten and
Loffler, 1990). For each of these examples, the differentiated
cells turn over rapidly and homeostatic mechanisms act on stem cells to
generate new cells. This ability to replenish certain single cell types is
crucial for maintaining the normal physiology and lifespan in humans. For
instance, the deficient replenishment of erythrocytes results in anemia,
whereas excessive turnover or misregulation of epidermal cells may cause
psoriasis or skin cancer (Weinstein and
Frost, 1968). Other human diseases result from deficits in single
cell types with less capacity for regeneration, such as pancreatic β
cells in autoimmune type I diabetes, or oligodendrocytes in multiple sclerosis
(for reviews, see Bach, 1994;
Compston and Coles, 2002). An
understanding of the mechanisms regulating single cell type regeneration in
model systems might therefore be used to develop cures for relevant human
diseases.

An attractive cell type for this approach is the zebrafish melanocyte,
because it is easily visualized, dispensable for viability in the laboratory,
and multiple mutations have been generated that affect different aspects of
its development at different stages of the fish life cycle. As in other
vertebrates, zebrafish melanocytes are derived from the embryonic neural crest
(Raible et al., 1992).
Melanocytes first appear at 24 hours postfertilization (hpf), and, by
approximately 60 hpf, the larval pigment pattern is established with
approximately 460 post-mitotic melanocytes. This number of larval melanocytes
remains nearly constant, with only minimal birth and death of melanocytes
occurring, until the onset of adult pigment pattern metamorphosis at 14 days
postfertilization (dpf) (Milos and Dingle,
1978; Johnson et al.,
1995) (reviewed by Rawls et
al., 2001). Previously, we demonstrated the homeostatic regulation
of larval melanocytes by larval melanocyte regeneration following melanocyte
ablation with a dermatology laser. We further revealed that this melanocyte
regeneration is achieved through the recruitment of undifferentiated cells,
which tends to exclude division of differentiated melanocytes (i.e.
compensatory regeneration, Fig.
1B) as the mechanism for larval melanocyte regeneration
(Yang et al., 2004). Despite
the power of such laser ablation, our attempts to further investigate the
mechanisms of larval melanocyte regeneration were hampered by the limiting
number of regenerating melanocytes that could be generated for thorough
analyses of cell division events.

Four possible modes for melanocyte regeneration distinguished by roles
for cell division and the differentiation state of precursors. (A)
Regeneration involves direct proliferation of pre-existing, differentiated
melanocytes and requires no further role for differentiation (compensatory
regeneration). (B) Regeneration is achieved through the recruitment of
quiescent precursors or stem cells to proliferate and differentiate
(epimorphosis). (C) Regeneration involves transdifferentiation of other
differentiated cells without cell division to generate the new melanocytes
(morphallaxis). (D) Regeneration occurs by direct differentiation of
set-aside late-stage undifferentiated precursors (for instance,
dct+ melanoblasts) into melanocytes without cell division.
Note that unlike stem cell-based models, regeneration by this last mechanism
would tend to exhaust the precursor pool.

In this study, we have developed a pharmacological method to ablate the
entire melanocyte population in zebrafish larvae. This now allows us to
further test the mechanisms underlying melanocyte regeneration. We show that a
small molecule, (2-morpholinobutyl)-4-thiophenol (MoTP), specifically ablates
melanocytes or melanoblasts, and that the melanocytotoxicity is mediated via
tyrosinase activity, presumably to convert MoTP to cytotoxic quinone species.
Following MoTP treatment, we demonstrate by BrdU incorporation experiments
that regenerated melanocytes are derived from the division of otherwise
quiescent melanocyte precursors or stem cells. We further show that this
regeneration requires the kit receptor tyrosine kinase. Together with
our analyses of melanocyte regeneration in wild-type and
kitj1e99 animals, we suggest that, in addition to
establishing the larval melanocyte pattern, embryos set aside a small number
of reserve cells (melanocyte precursors or stem cells) with an uneven
distribution. These cells can be drawn upon to re-enter developmental
pathways, divide multiple times and reconstitute the larval melanocyte
population following melanocyte ablation by MoTP treatment.

MATERIALS AND METHODS

Stocks

Zebrafish were reared according to standard protocols at 28.5°C, unless
otherwise noted (Westerfield,
1993). All developmental staging is reported in hours and days
postfertilization (hpf and dpf, respectively), corresponding to staging at the
standard temperature of 28.5°C. Staging at permissive temperature
(25°C) is translated to 28.5°C stages as previously described
(Kimmel et al., 1995).
kitj1e99 and fmsj4e1 have been
previously described (Rawls and Johnson,
2003; Parichy et al.,
2000). sdyj9s1 is a spontaneous, homozygous
viable allele of the shady locus
(Kelsh et al., 1996) that has
approximately 10% of the normal number of embryonic or larval iridophores
(C.-T.Y., unpublished). All references to these mutant alleles correspond to
homozygous animals.

Chemicals

(2-morpholinobutyl)-4-thiophenol (MoTP) was custom synthesized by Gateway
Chemical Technology (St Louis, MO). 4-hydroxyanisole (4-HA) (or
4-methoxyphenol) was purchased from Sigma-Aldrich (M1865-5, St Louis, MO). The
dose responses of both chemicals in melanocytotoxicity and fish lethality were
tested, and the most melanocytotoxic concentrations with low fish lethality
were chosen for zebrafish melanocyte ablation experiments. MoTP and 4-HA were
dissolved in dimethyl sulfoxide (DMSO) to make stock solutions, which were
then diluted in egg water at 14 μg/ml (50 μM) and 2 μg/ml (16 μM)
final concentrations, respectively. For phenylthiourea (PTU) treatment,
0.1-0.2 mM PTU was added to egg water and changed every 2 days
(Milos and Dingle, 1978).

BrdU incorporation and detection

To detect and quantify cell division events in melanocyte lineages during
larval melanocyte regeneration, normally reared and MoTP-treated larvae at
various developmental stages were immersed in a solution of
5-bromo-2′-deoxyuridine (BrdU; 5 mM) for 24 hours in pulse labeling
experiments. In the continuous labeling experiment, we changed the BrdU
solution every 24 hrs. At the harvest stages, animals were fixed, imbedded in
paraffin wax, and then cut into series of 5-μm-thick sagittal sections.
Sections were deparaffinized and incubated with mouse anti-BrdU monoclonal
antibody (1:300 dilution, Santa Cruz), followed by a secondary incubation with
Alexa Fluor 594-conjugated goat anti-mouse Ig (1:300 dilution, Molecular
Probes), and bisbenzimide (100 ng/ml; Sigma) for nuclear staining
(Rawls et al., 2004). BrdU
incorporation was determined by first identifying bisbenzimide-positive nuclei
(blue fluorescence) in pigmented melanocytes, then assessing BrdU
incorporation (red fluorescence) in each identified nucleus
(Fig. 5).

RESULTS

Melanocyte differentiation proceeds to tyr+ and
dct+ stages in the presence of MoTP

To identify a chemical that ablates the entire larval melanocyte
population, we sought to investigate the biological effect of MoTP in
zebrafish melanocyte development. MoTP was first described in a small molecule
screen for drugs that affect zebrafish development
(Peterson et al., 2000). These
researchers observed that embryos developing in the presence of MoTP lacked
body melanocytes, whereas their retinal pigment epithelium (RPE) was lightly
pigmented. This differential effect on RPE and neural crest (NC)-derived
melanocytes suggests that MoTP does not merely block general melanin
production, for instance as does a tyrosinase inhibitor such as PTU.
Furthermore, Peterson et al. reported that when larvae were removed from MoTP
after a three-day incubation, melanocytes gradually repopulated the larval
body during the ensuing two days. Together, these observations led to the
suggestion that MoTP reversibly blocks or delays NC melanocyte development in
zebrafish (Peterson et al.,
2000).

To test the hypothesis that MoTP causes developmental delay in the
melanoblast lineages, we examined their development in MoTP-treated larvae by
performing a series of whole-mount RNA in situ hybridization with the early
stage melanoblast markers mitf
(Lister et al., 1999) and
kit (Parichy et al.,
1999), and the late stage melanoblast markers tyr
(Page-McCaw et al., 2004) and
dct (Kelsh et al.,
2000) (C.-T.Y., unpublished). For these experiments, we incubated
14 hpf embryos in 14 μg/ml MoTP solution and fixed them at 27 hpf for in
situ analysis. Following in situ hybridization, we were unable to distinguish
differences in the developmental patterns or numbers of labeled cells in
MoTP-treated larvae compared with untreated larvae for each of the markers
(Fig. 2A-H). These results
indicate that the differentiation of NC-derived melanoblasts remains normal
and proceeds to late developmental stages (tyr+ or
dct+) in the presence of MoTP. Interestingly, untreated
late-stage melanoblasts typically begin to melanize prior to 27 hpf, but in
MoTP treated larvae, this terminal marker (melanin) fails to form.

MoTP is cytotoxic to larval melanocytes

Because our foregoing analysis revealed that MoTP-treated melanoblasts
proceed to late stages of differentiation but fail to reach their terminal
stage of melanin production, we next explored their fate at subsequent
developmental periods in MoTP-treated larvae. We found that the number of
dct+ cells in treated larvae declined from a maximum of
100 at 31 hpf to an average of 15 dct+ cells by 60 hpf
(Fig. 3B,D,F-H). By contrast,
in the untreated larvae, the number of dct+ cells steadily
increased to more than 300 during the same time period
(Fig. 3A,C,E,H). Note that the
majority of the dct+ cells in the untreated larvae are
differentiated melanocytes, whose subsequent melanin production was blocked by
incubation with PTU, allowing for easier visualization of the in situ marker
(see Materials and methods). The disappearance of dct+
cells in treated larvae raised the possibility that, in the presence of MoTP,
melanoblasts reach the tyr and dct expression stage, at
which time they become sensitive to MoTP and die.

Melanocyte lineages develop to late stages in the presence of MoTP.
(A-H) The development of melanocyte lineages in MoTP-treated (B,D,F,H)
and untreated (A,C,E,G) larvae were examined by whole-mount RNA in situ
hybridization with mitf (A,B), kit (C,D), tyr (E,F)
and dct (G,H). For these experiments, embryos were incubated in MoTP
solution from 14 to 27 hpf, then immediately fixed for in situ analysis. The
developmental patterns and the numbers of cells (white arrowheads) for each of
the markers are indistinguishable in the MoTP-treated larvae from those in
untreated larvae. Untreated larvae were reared in PTU to completely block
melanin synthesis, thereby allowing easier visualization of NBT/BCIP
precipitates in melanocytes. Scale bar: 500 μm.

We explored the notion that MoTP ablates cells in the melanocyte lineages
by observing its effects on melanocytes that were first allowed to melanize in
the absence of MoTP. When 48 hpf larvae with pigmented melanocytes were
incubated in MoTP solution, larval melanocytes became punctate within 12 hours
(Fig. 4A-D), and, by 24 hours,
these punctate melanocytes began to extrude from the epidermis
(Fig. 4E). As these two
phenotypes are hallmarks of melanocyte cell death in fish
(Parichy et al., 1999;
Sugimoto et al., 2000), we
interpret these results to indicate that MoTP induces melanocyte cell
death.

The melanocytotoxicity of MoTP is dependent on tyrosinase
activity

We sought to investigate the mechanism underlying MoTP cytotoxicity in
melanocytes. One class of phenolic compounds has been previously reported to
cause cytotoxicity specific to melanocytes (melanocytotoxicity) in mammals
(reviewed by Riley, 1985).
These compounds feature a phenolic ring with a functional group at the para
position, a structure that is similar to tyrosine, the initial substrate of
tyrosinase during the biochemical synthesis of melanin. Biochemical studies
have suggested that this class of phenolic compounds, such as 4-hydroxyanisole
(4-HA; Fig. 4G), competes with
tyrosine for hydroxylation by tyrosinase and, consequently, is converted to a
cytotoxic form, mainly o-quinone, which is associated with the initiation of
cellular damage in melanocytes (Riley,
1975; Naish et al.,
1988). Thus, these studies demonstrated that the
melanocytotoxicity of such phenolic compounds is mediated by tyrosinase
activity (Fig. 4I). Like known
members of this class of phenolic compounds, MoTP also has a phenolic ring
with a functional group, in this case (morpholinobutyl)-thio, at the para
position. Therefore, we sought to test whether MoTP acts similarly to the
4-HA-related class of phenolic compounds. If so, we reasoned that the
chelation of copper, an essential cofactor for tyrosinase activity, by
addition of the copper chelator PTU, should block the melanocytotoxicity of
MoTP (Fig. 4I). Accordingly, we
co-incubated 48 hpf larvae in MoTP solution with 0.1-0.2 mM PTU. We found that
the melanocytotoxicity of MoTP was completely suppressed in the presence of
PTU, as lightly pigmented and dendritic melanocytes were observed, but we
observed no evidence of punctate melanocytes or their subsequent extrusions in
these larvae (Fig. 4F).

The number of dct+ cells declines in the presence of
MoTP. (A-H) The development of late-stage dct+
cells was further examined as the MoTP incubation continued. Larvae were
reared in MoTP solution beginning at 14 hpf and fixed at selected
developmental stages up until 68 hpf for in situ analysis. (A,C,E) The
distribution and the number of dct+ cells in untreated
larvae (reared in PTU solution) at 37, 42 and 61 hpf, respectively. Note that
many dct+ cells in the untreated animals are
differentiated melanocytes that remain unpigmented as a result of PTU
treatment. (B,D,F) dct+ cells in the MoTP-treated larvae
at 37, 42 and 61 hpf, respectively, revealing a gradual disappearance of
dct+ cells during the MoTP incubation. The number of
dct+ cells in the MoTP-treated larvae begins to decline
from a maximum of ∼100 dct+ cells at 31 hpf to an
average of 15 dct+ cells by 60 hpf (red line in H). These
dct+ cells have a stereotyped spacing pattern in the
dorsum (white arrowheads in G). By contrast, the number of
dct+ cells steadily increases to more than 300 in the
untreated larvae (blue line in H). Scale bar: in B, 500 μm for A-D; in F,
400 μm for E,F; in G, 400 μm.

The melanocytotoxicity of MoTP is mediated by tyrosinase activity.
(A) The chemical structure of (2-morpholinobutyl)-4-thiophenol (MoTP).
(B) Dendritic (healthy) melanocytes (black arrowheads) in an untreated
larva at 3 dpf. (C) When larvae were incubated in MoTP solution from 14
to 72 hpf, no neural crest-derived melanocytes were observed, but RPE is
lightly pigmented (black arrow). (D,E) When 48 hpf larvae with
pigmented melanocytes were shifted to MoTP, within 24 hours, larval
melanocytes had become punctate (white arrowheads in D) and began to extrude
from the epidermis (white arrow in E), a hallmark of melanocyte cell death.
(F) The melanocytotoxicity of MoTP was blocked by PTU, as indicated by
the dendritic appearance of lightly pigmented melanocytes (black arrowheads in
F) following co-incubation of MoTP and PTU from 48 to 72 hpf. (G) The
chemical structure of 4-hydroxyanisole (4-HA). (H) When larvae were
incubated with 4-HA from 14 to 72 hpf, the same punctate melanocyte pattern of
cell death appeared (white arrowheads). (I) Illustration of the
mechanism of 4-HA melanocytotoxicity (see
Riley, 1985). Note that
tyrosinase converts the prodrug 4-HA to a cytotoxic o-quinone. Because copper
is an essential cofactor for tyrosinase, its activity is blocked by
co-incubation with PTU, a copper chelator. Timelines (gray) below the panels
indicate the period of drug treatments (red) and analysis time (vertical line
above timelines). Scale bars: in H, 500 μm for B-D,F,H; in E, 50 μm.

To further demonstrate that the melanocyte cell death caused by MoTP is
related to the melanocytotoxicity described for 4-HA, we tested the effects of
4-HA in zebrafish. When embryos were reared in 2 μg/ml 4-HA solution (14-72
hpf), the same punctate melanocyte patterns of cell death were observed as we
described following MoTP treatment (Fig.
4H). In addition, the melanocytotoxicity of 4-HA was abrogated by
co-incubation with PTU (data not shown). These results suggest that MoTP acts
by the same, or a similar, mechanism as 4-HA conversion from a prodrug into a
cytotoxic form by the activity of tyrosinase, an enzyme found at high levels
in melanocytes.

Melanocyte regeneration following MoTP-induced ablation

As described above, incubating zebrafish embryos in MoTP results in the
loss of melanoblasts or melanocytes. By 72 hpf, when untreated larvae have
established the larval melanocyte pattern, MoTP-treated larvae lack all
melanocytes. When MoTP-treated larvae are transferred to fresh egg water at 72
hpf, melanocytes gradually develop over the next 4-5 days (as described in the
ensuing sections). Such regeneration of larval melanocytes now provides an
opportunity to explore how regenerating cells are derived from their
precursors, including the role for cell division and differentiation state of
precursors (below).

Melanocytes regenerate from the division of undifferentiated
melanocyte precursors following MoTP treatment

We first sought to test whether, following melanocyte ablation by MoTP,
melanocytes regenerate through mechanisms involving cell division.
Accordingly, we incubated larvae in MoTP from 4-5 dpf, to ablate
differentiated melanocytes. We also incubated these larvae continuously in
BrdU from 4-10 dpf, to reveal whether the differentiated melanocytes that
arise have undergone rounds of DNA synthesis, BrdU incorporation, and thus,
cell division. Counting the percentage of BrdU-positive, melanin-positive
melanocytes in 5 μm paraffin sagittal sections at 10 dpf (5 days post-MoTP
treatment) showed that 97.2±2.5% of regenerated melanocytes are BrdU
positive, compared with only 3.9±3.5% BrdU-positive melanocytes in
untreated animals (Fig. 5).
This finding indicates that virtually all regenerated melanocytes develop from
precursors that have undergone one or more rounds of cell division following
MoTP-induced melanocyte ablation.

We next explored models of regeneration that do not involve cell division
(Fig. 1C,D). We reasoned that
if the embryo sets aside late-stage precursors that could differentiate
directly into melanocytes without cell division for regeneration and
regulation (Fig. 1D), these
late stage precursors might express a late-stage melanoblast marker, such as
dct, in untreated (non-regenerating) larvae. In situ analysis of
dct expression showed very few dorsal dct+
melanoblasts (0.2-0.3 per animal) in untreated fish between 74 and 128 hpf
(Fig. 6C). By contrast, in
MoTP-treated larvae (14-72 hpf), we detected 8- to 20-fold more dorsal
dct+ melanoblasts (two to three per animal) at the same
developmental stages (Fig.
6A-C). This result indicates that large numbers of late-stage
(dct+) melanoblasts are not available for regenerating the
larval melanocyte pattern, and also that ablation of the embryonic
melanoblasts and melanocytes results in the recruitment of
dct+ melanoblasts from less-differentiated precursors
during melanocyte regeneration.

An alternative model of regeneration without cell division is regeneration
by transdifferentiation from one differentiated cell type to another. For
example, in amphibians, other NC-derived chromatophores have been suggested to
transdifferentiate into melanophores under various experimental conditions
(Thidaudeau and Holder, 1998; Ide and
Akira, 1988); thus, differentiated iridophores and xanthophores
may be candidates for transdifferentiation into melanocytes in this
regeneration system (Fig. 1C).
Evidence for or against such transdifferentiation might be gained by assessing
regeneration in the absence of iridophores or xanthophores. Accordingly, we
investigated whether melanocyte regeneration occurred in mutant larvae that
lacked most or all iridophores (sdyj9s1) or xanthophores
(fmsj4e1), but that have a normal embryonic and larval
melanocyte pattern. In each mutant, following melanocyte ablation by MoTP
treatment (14-72 hpf), we observed that melanocyte regeneration occurred
identically to that in wild-type larvae (data not shown), which suggests that
transdifferentiation from iridophores and xanthophores is not responsible for
melanocyte regeneration. We note, however, that these results do not exclude
the possibility of transdifferentiation from other cell types.

Larval melanocyte regeneration following MoTP treatment is achieved by
cell division. (A-H) Cell division events in melanocyte lineages
during larval melanocyte regeneration were tracked by BrdU incorporation
experiments. Larvae were continuously incubated in BrdU (5 mM) during and
after MoTP treatment, then fixed, paraffin embedded and 5 μm sagittal
sections processed for BrdU immunohistochemistry after melanocyte regeneration
was mostly completed at 10 dpf (5 days post-MoTP treatment). BrdU
incorporation states of pigmented melanocytes were assessed by first
identifying melanocyte nuclei (thinning in melanin, white arrowheads in A and
E) accompanied by bisbenzimide staining (white arrowheads in B and F). These
were then examined for red fluorescence indicative of BrdU incorporation
(white arrowheads in C and G). D is the overlay image of A, B and C,
indicating a melanocyte that did not incorporate BrdU during the BrdU
labeling, whereas in H, the overlay image of D, E and F shows a different
melanocyte that did incorporate BrdU during the BrdU labeling. (I)
Quantitative analyses of BrdU incorporation in larvae exposed to MoTP from 4
to 5 dpf. Black and white bars indicate BrdU incorporation in untreated and
MoTP-treated larvae, respectively. Horizontal green line indicates periods of
BrdU labeling. Asterisk in I indicates the developmental stages at which
larvae were sacrificed for BrdU incorporation analysis. Scale bar in A: 10μ
m for A-H.

Taken together, these results suggest that regenerated melanocytes arise by
the recruitment of undifferentiated melanocyte precursors that then divide
several times and differentiate into new melanocytes. Our results tend to
exclude possible regeneration from differentiated melanocytes
(Yang et al., 2004),
iridophores or xanthophores, or from late-stage (dct+)
precursors.

Appearance of dct+ melanoblasts following
MoTP-induced melanocyte death. (A,B) Timeline for A and B is
shown below A. Lightly pigmented, new melanocytes (black arrow; A,B) and
dct+ melanoblasts (white arrows, B) appear 24-36 hours
postMoTP treatment. (C) The appearance of the dct+
melanoblasts in the dorsal stripe is specifically induced in the MoTP-treated
larvae, average of two to three dct+ melanoblasts per
animal examined (white bars), compared with less than 0.3
dct+ melanoblasts (black bars) at equivalent stages in
untreated larvae. Eight to 18 animals were analyzed for each bar. Scale bars:
in A, 500 μm; in B, 100 μm.

Dynamics of regenerated melanocyte precursor cell division

We were interested in the dynamics of the cell division during larval
melanocyte regeneration. Accordingly, following MoTP incubation (14-72 hpf) to
ablate the ontogenetic melanocyte population, we monitored the history of cell
division by pulse-labeling regenerated melanocytes with BrdU for 24 hours at
various regeneration stages, and assessed the percentage of BrdU-positive
melanocytes at 8 dpf (5 days post-MoTP treatment). We found that approximately
35-40% of melanocytes incorporated BrdU on each of the second and third days
post-MoTP treatment. The percentage of melanocytes that had incorporated BrdU
during the labeling period then declines to approximately 20% by the fourth
and fifth days post-MoTP treatment. By contrast, we found that only 5-8% of
melanocytes in the untreated larvae incorporated BrdU at the equivalent
developmental stages (Fig. 7A).
These results suggest that cell divisions in melanocyte precursors occur
continuously throughout the 5 days of regeneration.

Dynamics of cell divisions during melanocyte regeneration.
(A,B) The history of cell divisions in melanocyte lineages
during larval melanocyte regeneration was determined by pulse-labeling with
BrdU for 24 hours at various stages during or after MoTP treatment.
Quantitative analyses of BrdU incorporation were conducted in larvae exposed
to MoTP from 14 to 72 hpf (A), or 4 to 5 dpf (B). Black and white bars
indicate BrdU incorporation in untreated and MoTP-treated larvae,
respectively. Horizontal green lines indicate periods of BrdU labeling.
Asterisks in A and B indicate the developmental stages at which larvae were
sacrificed for BrdU incorporation analysis.

We further investigated how rapidly melanocyte precursors or stem cells are
induced to re-enter the cell cycle upon melanocyte ablation by MoTP treatment.
Because we found that we could not achieve reliable BrdU incorporation in
zebrafish larvae by simply rearing larvae in BrdU solution prior to 4 dpf, we
instead used a later stage for MoTP melanocyte ablation. Accordingly, we
ablated larval melanocytes by MoTP treatment from 4 to 5 dpf, incubated larvae
with BrdU during the 4 to 5 and 5 to 6 dpf periods, and then assessed BrdU
incorporation in melanized melanocytes after regeneration was near completion
at 9 dpf. We found that 47% of melanocytes had incorporated BrdU during the 4
to 5 dpf labeling period (coincident with the MoTP treatment), rising to 56%
BrdU-positive melanocytes during the 5 to 6 dpf period
(Fig. 7B). These results
indicate that melanocyte precursors or stem cells are induced to enter the
cell cycle within 24 hours of the beginning of MoTP incubation. Because signs
of melanocyte cell death occur as early as 12 hours after the beginning of
MoTP exposure, we suggest that this induction may begin less than 12 hours
after melanocyte cell death.

Recruitment of melanocyte precursors or stem cells in wild-type and
mutant animals

We were also interested in the distribution of quiescent melanocyte
precursors or stem cells that contribute to melanocyte regeneration following
melanocyte ablation by MoTP treatment. Ideally, this knowledge could be gained
by examining markers for the melanocyte stem cells. Lacking such stem cell
markers, we have instead examined the distribution of differentiated
melanocytes in wild-type and mutant animals for less direct inferences. Thus,
in wild-type animals, we observed the first appearance of regenerated
melanocytes 24 hours post-MoTP treatment. The number of melanocytes continues
to steadily rise for the ensuing 4 days and reaches a plateau of approximately
350-400 melanocytes by 9 dpf (6 days after the removal of MoTP). This number
reveals a deficit of approximately 60-110 melanocytes in MoTP-treated versus
untreated larvae (average 460 melanocytes;
Fig. 8A). To further explore
this deficit, we examined the patterns of melanocytes in the MoTP-treated and
untreated larvae at 9 dpf. We found MoTP-treated larvae regenerate an almost
identical pigment pattern compared to that of the untreated larvae, with a
similar number and distribution of melanocytes in the dorsal, lateral and
ventral larval melanocyte stripes. One exception was that MoTP-treated larvae
failed to regenerate the majority of the ventral yolk sac melanocytes
(Fig. 8B-E). Typically,
untreated larvae have approximately 65 melanocytes in the ventral yolk sac
stripe at this stage; therefore, the absence of those melanocytes in
MoTP-treated larvae accounts for the observed melanocyte deficit
(Fig. 8A). When we allowed
these MoTP-treated larvae to develop to stages immediately prior to the onset
of adult pigment pattern metamorphosis, we observed that melanocytes slowly
begin to repopulate the ventral yolk sac area, resulting in approximately 35%
of the normal number of ventral yolk sac stripe melanocytes by 14 dpf (11 days
post-MoTP treatment; data not shown). These results suggest either that the
region of the ventral yolk sac is devoid of quiescent melanocyte precursors or
stem cells, or that they are less responsive to the presence or absence of
differentiated melanocytes in this area (see Discussion).

Ventral yolk sac melanocytes fail to regenerate following melanocyte
ablation by MoTP treatment. (A) Larval melanocytes were counted in
the untreated larvae (blue line) and MoTP-treated (14-72 hpf) larvae (red
line). After the appearance of melanocytes at 24 hours post-MoTP treatment,
the number of melanocytes continues to steadily rise in the ensuing 4 days and
then plateaus at approximately 350-400 melanocytes by 9 dpf (6 days after
removal of MoTP). (B-E) At this stage of melanocyte regeneration,
MoTP-treated larvae (C,E) have regenerated almost identical pigment patterns
to those of the untreated larvae (B,D), with a similar number and distribution
of melanocytes in the dorsal, lateral and ventral larval melanocyte stripes
(white arrowheads). However, MoTP-treated larvae fail to regenerate the
majority of the ventral yolk sac melanocytes (white arrow in C and E). Scale
bars: 500 μm.

To further understand the mechanisms by which melanocyte homeostasis is
regulated by quiescent melanocyte precursors or stem cells, we examined
melanocyte regeneration in mutants for kit, which has been suggested
to play important roles in the recruitment of melanocyte stem cells or
expansion of melanoblasts (Rawls and
Johnson, 2001). The kit receptor tyrosine kinase is
required for ontogenetic melanocyte development in zebrafish embryos. In
kitb5 (a null mutation) embryos, NC-derived melanocytes
differentiate, but fail to migrate to proper locations and subsequently
undergo programmed cell death, resulting in larvae devoid of melanocytes until
pigment pattern metamorphosis (Johnson et
al., 1995; Parichy et al.,
1999). Previously, by using a temperature-sensitive allele,
kitj1e99, at the restrictive temperature (30-33°C), we
showed that kit is also required to fill in gaps in the larval
melanocyte pattern created by laser ablation. Interestingly,
kitj1e99 animals also failed to regenerate melanocytes
after laser ablation at temperatures described as permissive for embryonic
melanocyte development or for melanocyte regeneration in the regenerating
adult caudal fin (23 or 25°C) (Yang et
al., 2004; Rawls and Johnson,
2001; Rawls and Johnson,
2003). Because the laser ablations resulted in animals with only
small regions devoid of melanocytes, we revisited the role of
kitj1e99 in larval melanocyte regeneration using MoTP
treatment to ablate all the melanocytes in the larvae. When we reared
kitj1e99 animals in MoTP solution (14-72 hpf) at the
permissive temperature (25°C), we found that new melanocytes appeared
significantly more slowly than in wild-type larvae after MoTP treatment
(Fig. 8A,
Fig. 9A), achieving a plateau
of 15.9±5.7 melanocytes at 9 dpf. We discuss the inferences that can be
drawn from the number and distribution of these few regenerated melanocytes
more extensively below (see Discussion).

DISCUSSION

Zebrafish melanocyte ablation by MoTP

To complement the traditional regeneration studies of vertebrate appendages
or organs, we have developed methods to ablate a single cell type, the
melanocyte, and analyze its regeneration in zebrafish larvae. Previously, we
have shown that a standard, hand-held dermatology laser can be used to
specifically ablate melanocytes from small regions of zebrafish larvae,
allowing us to study melanocyte regeneration in otherwise intact animals
(Yang et al., 2004). Here, we
have developed a different method, which can ablate the entire melanocyte
population in zebrafish larvae. We demonstrated that a small molecule, MoTP,
is cytotoxic to late stage melanoblasts and melanocytes in zebrafish larvae,
and that this melanocytotoxicity is dependent on tyrosinase activity. The
melanocyte specificity of MoTP ablation is also stage specific; we found that
exposure of larvae to MoTP after 5 dpf had no effect on melanocytes (data not
shown), presumably because of the decline of tyrosinase activity in mature
melanocytes. Although tyrosinase is also expressed in the RPE
(Camp and Lardelli, 2001), we
found that the RPE is relatively insensitive to MoTP-induced cell death, as
the RPE develops normally, albeit with lighter pigment, at the MoTP
concentration that causes NC-derived melanocyte cell death
(Fig. 4C). This differential
sensitivity between types of melanocyte could result from differential
tyrosinase expression or activity, or possibly from differences in the
accessibility of these cells to MoTP during the treatment.

A similar melanocytotoxicity has been described for a related class of
compound, including 4-HA. Like MoTP, 4-HA is a prodrug that is converted by
tyrosinase to a cytotoxin. In the case of 4-HA, the cytotoxic product is
o-quinone (Naish et al.,
1988). Thus, we predict that tyrosinase converts MoTP to a similar
molecule. The melanocytotoxicity of 4-HA was explored for use in melanoma
chemotherapy, but discontinued because of severe liver damage resulting from
liver cytochrome P450 conversion of 4-HA to toxic p-quinone
(Rustin et al., 1992;
Moridani et al., 2002). The
difference in structure of MoTP from 4-HA in the functional group at the para
position raises the possibility that MoTP or related derivatives may not show
the same liver toxicity. The recent development of a melanoma model in
zebrafish may provide a useful platform for evaluating MoTP as a
chemotherapeutic, or for the search for other tyrosinase-dependent prodrugs
for the treatment of melanoma (Patton et
al., 2005).

In addition, the finding that MoTP is a prodrug converted by tyrosinase to
a cytotoxic form suggests the potential application that specific and
non-invasive ablation of any cell type in complex tissues or organs can be
achieved by expressing tyrosinase under the control of cell- or
tissue-specific promoters, then exposing the fish to MoTP. A similar use for
transgenic expression of a prodrug converting enzyme, which we are exploring,
is expression of bacterial nitroreductase in specific cells
(Medico et al., 2001) (M. T.
Saxena and S. L. Johnson, unpublished), followed by exposure to bacterial
enzyme-specific substrates metronidazole or CB1954, that in turn are converted
to cytotoxins.

We note that Harris et al. (Harris et
al., 2003) have previously demonstrated an example of single cell
type ablation by small molecules in zebrafish; aminoglycoside antibiotics,
including neomycin, can induce hair cell death in larval zebrafish lateral
line neuromasts, which is then followed by hair cell regeneration.

We explored the developmental mechanisms by which melanocytes regenerate
following melanocyte ablation by MoTP treatment, particularly in regards to
the role of cell division and the differentiated state of the regeneration
precursor cells (Fig. 1). The
finding that melanocyte regeneration occurs after essentially complete
ablation of the larval melanocytes by MoTP, and our previous findings
(Yang et al., 2004) that
larval melanocytes regenerate from undifferentiated precursors, rather than
differentiated melanocytes, following laser ablation, rule out that melanocyte
regeneration is achieved via the proliferation of differentiated melanocytes
(Fig. 1A). This finding,
together with our finding that all or virtually all regenerated melanocytes
arise via cell division after ontogenetic melanocyte ablation
(Fig. 5I), provides strong
support for the model outlined in Fig.
1B that larval melanocyte regeneration occurs via the recruitment
of undifferentiated precursors to divide and then differentiate to produce the
new larval melanocyte population (an example of epimorphosis).

Our findings also provide evidence against models of regeneration that do
not involve cell division (Fig.
1C,D). The notion that late stage (dct+)
precursors are available to differentiate directly into melanocytes without
cell division (Fig. 1D) is
dispelled by our finding of few dct+ melanoblasts in
untreated larvae, especially when compared with the numbers of
dct+ melanoblasts found in MoTP-treated larvae
(Fig. 6C). The notion of direct
transdifferentiation (Fig. 1C)
from other pigment cells playing a role in melanocyte regeneration is
suggested by findings of transdifferentiation between chromotaphores in
salamanders (Thidaudeau and Holder, 1998;
Ide and Akira, 1988). However,
our finding that sdy and fms mutants that lack most or all
iridophores or xanthophores, respectively, regenerate melanocytes identically
to wild-type larvae does not support the notion that melanocyte regeneration
is derived from these other pigment cell types.

Melanocyte regeneration deficit in kitj1e99
larvae. (A) Larval melanocytes were counted in the untreated (blue
line) and MoTP-treated (14-72 hpf; red line) kitj1e99
larvae at the permissive temperature (25°C) for the temperature-sensitive
allele. The first few melanocytes appear approximately 3 days post-MoTP
treatment in the MoTP-treated kitj1e99 animals at
25°C. The number of melanocytes in MoTP-treated larvae reaches
approximately 15 by 9 dpf (6 days after removal of MoTP, red line in A),
whereas age-matched untreated kitj1e99 animals have
approximately 350-400 melanocytes (blue line in A). (B,C) Note
that the regenerated melanocytes exhibit a stereotyped spacing morphology in
the dorsum (white arrowheads in C). B and C are untreated and MoTP-treated
kitj1e99 larvae at 7 dpf, respectively. Scale bar: 500μ
m

These studies show that melanocytes in larval zebrafish regenerate from
undifferentiated precursors that may undergo several rounds of cell divisions
prior to differentiation to replace the missing melanocyte pattern. It will be
interesting to discover how much of this mechanism of recruiting
undifferentiated precursors to reenter developmental pathways in this
regeneration system is shared by the regulatory mechanism that acts in the
embryo to precisely determine melanocyte number.

Melanocyte stem cells in zebrafish larvae

Homeostasis or regeneration of many tissues has been shown to be achieved
via the recruitment of stem cells. Stem cells have two important
characteristics: they are undifferentiated and they self-renew
(Siminovitch et al., 1963).
These two key characteristics of stem cells were elegantly demonstrated by
multiple reconstitutions of hematopoietic cells in X-ray-irradiated mice
(Spangrude et al., 1988).
Because such techniques of transplantation and reconstitution are not
generally possible for other lineages, other lines of evidence may help
evaluate the notion of stem cells or quiescent precursors. The presence of
melanocyte stem cells in zebrafish has been suggested by observations of the
unlimited capacity of melanocyte pattern re-establishment in the regenerating
caudal fin (Rawls and Johnson,
2000). Melanocyte or pigment cell stem cells have also been
suggested to contribute to the adult melanocyte population during
metamorphosis, indicating that stem cells are established during embryonic or
larval stages (Johnson et al.,
1995; Parichy,
2003). Assuming that each of these events draws upon the same
precursor population, our finding of a high proliferative capacity of the
melanocyte precursors during larval melanocyte regeneration suggests that
these melanocyte precursors have many of the characteristics expected of stem
cells. Finally, the demonstration of melanocyte stem cells in mammalian hair
follicles (Nishimura et al.,
2002) supports the idea of an analogous stem cell in the
zebrafish.

Zebrafish melanocytes begin to appear at 24 hpf, and the larval pigment
pattern is established at approximately 60 hpf. Our results suggest that in
addition to establishing the larval melanocyte pattern, embryos set aside a
population of quiescent reserve cells, henceforth melanocyte stem cells, that
can be drawn upon to generate new melanocytes for larval melanocyte
homeostasis; for instance, for filling gaps in the melanocyte pattern
(Yang et al., 2004), or for
later stages of melanocyte development, such as for adult pigment pattern
formation (Johnson et al.,
1995; Parichy,
2003). Our BrdU incorporation experiments suggest that melanocyte
stem cells are maintained in a minimal state of activity during the larval
stage. Upon melanocyte ablation by MoTP treatment, melanocyte stem cells are
released from this quiescent state, divide and produce new melanocytes to
replenish the ablated population. The activity of these melanocyte stem cells
and their descendant amplifying melanoblasts appears to be highly regulated
during melanocyte regeneration. Our BrdU incorporation experiments show that
the recruitment of melanocyte stem cells into the cell cycle is relatively
rapid, beginning less than 24 hours after melanocyte cell death. Furthermore,
we observe cell divisions throughout the 5-day regeneration period with fewer
cell divisions at the end of this period
(Fig. 7). The fact that the
pattern is largely regenerated with no local excesses or deficits of
melanocytes (with the exception of the ventral yolk sac stripe melanocytes,
see discussion below) also suggests that feedback regulation from the final
pattern regulates the later cell divisions.

One intriguing finding in our regeneration studies is that, following
melanocyte ablation by MoTP treatment, ventral yolk sac stripe melanocytes
fail to regenerate (Fig. 8).
The melanocyte stripe on the ventral yolk sac is the last larval melanocyte
stripe established, and, furthermore, these melanocytes migrate the largest
distance from the dorsum to their final ventral locations. Uneven distribution
of melanocyte progenitors has been described to account for the varying
densities of melanoblasts in different parts of the mouse embryo (reviewed by
Besmer, 1993;
Wilkie et al., 2002).
Following this logic, the failure of larval melanocyte regeneration in the
zebrafish ventral yolk sac could be due to the lack of melanocyte stem cells
in the region. Alternatively, melanocyte stem cells may in fact be established
in the ventral yolk sac but fail to survey the environment for melanocyte
homeostasis, or to re-enter developmental pathways upon the melanocyte
ablation by MoTP treatment. Interestingly, one third of the normal number of
ventral yolk sac stripe melanocytes eventually reappears by late larval stage,
during the onset of adult pigment pattern metamorphosis (14 dpf, data not
shown). An attractive possibility is that this late stage regeneration now
takes advantage of melanocyte stem cells primed by the onset of pigment
pattern metamorphosis to reconstitute the previously ignored yolk sac
melanocyte deficit. Such models can be further tested once markers that label
melanocytes precursors or stem cells are developed.

An interesting question of melanocyte regeneration is how many melanocyte
stem cells in the zebrafish larvae contribute to the reconstitution of the
entire larval melanocyte population following melanocyte ablation. One
approach to answer this question is to estimate the number of cell divisions
that each melanocyte has progressed through during regeneration. Based on our
BrdU incorporation experiments, we estimate that melanocyte stem cells or the
descendant amplifying melanoblasts may have gone through as many as two to
four cell divisions. This is suggested in part by the accumulated percentage
of BrdU-incorporated melanocytes observed over the six intervals of 24-hour
BrdU labeling during and after MoTP incubation, indicating an average of 1.8
cell divisions for each melanocyte lineage
(Fig. 7). This number of cell
divisions per lineage is an underestimate, as some lineages are likely to have
divided twice or more during any one period of 24-hour BrdU labeling. In
addition, our observation that some melanocytes differentiate and leave the
cell cycle during the early stage of regeneration suggests that those that
differentiate at late stage of regeneration have undergone even more cell
divisions. From these calculations, we estimate that there are approximately
35-145 melanocyte stem cells in larval zebrafish that give rise to the
approximately 350-400 melanocytes present at the completion of
regeneration.

Another estimate for the number of stem cells comes from our observation of
dct-expressing cells during prolonged MoTP exposure. We typically
observe 15-20 dct+ cells in the larvae during prolonged
MoTP incubation (50-68 hpf; Fig.
3H). In addition, the dct+ cells on the dorsum
of these larvae are distributed with a stereotyped spacing pattern
(Fig. 3G). We suggest that
these cells are newly progressed to dct+ stage, and that
they then die once sufficient MoTP-induced cytotoxins accumulate. An
attractive possibility is that these dct+ cells each mark
the position or the domain of a single melanocyte precursor or melanocyte stem
cell, and the that number of these dct+ cells reflects the
number of melanocyte stem cells. Because, at any given time, some stem cell
lineages may not have dct+ daughters, the number of
melanocyte stem cells may be greater than the number of
dct+ cells present in a single fish at a particular
moment.

Our study of larval melanocyte regeneration in
kitj1e99animals at permissive temperature may provide
additional evidence for the above estimate of the number and position of
melanocyte stem cells. Among its roles in melanocyte development, kit
has been suggested to be required for melanoblast proliferation in mammals
(Mackenzie et al., 1997). In
zebrafish larvae, in addition to defects in melanocyte migration and
subsequent survival, kitb5 null mutants also develop a
reduced number of embryonic melanocytes (50-60% of the wild-type melanocyte
number), consistent with a role in proliferation as well
(Parichy et al., 1999). Here,
we find that following melanocyte ablation and recovery at the otherwise
permissive temperature for the allele, kitj1e99 animals
regenerate approximately 5% (15.9±5.7) of the melanocyte number
regenerated in wild-type animals (Fig.
9). The mutant lesion for this allele is in the second kinase
domain (Rawls and Johnson,
2003), and seems to confer temperature-sensitivity on the gene
product. The different effects of the kitj1e99 allele
between ontogeny and regeneration may reveal a regeneration-specific function
of the kit receptor tyrosine kinase mapping to this site, or,
alternatively, that the mutation partially reduces kit function at
the permissive temperature, and the regeneration role has a greater demand on
kit activity. Whichever is the case, the distribution of melanocytes
that regenerate in kitj1e99 animals may be informative of
the distribution of melanocyte precursors or stem cells. We note that
regenerated melanocytes in the dorsum of kitj1e99animals
are distributed in a similar pattern to the dct+ cells
observed in the prolonged MoTP-treated animals discussed above
(Fig. 3G,
Fig. 9C). One possibility is
that each melanocyte stem cell gives rise to a single melanoblast that
differentiates without further cell division. If this were so, then the number
of regenerated melanocytes observed in the kitj1e99animals
may directly reflect the number, and possibly the position, of melanocyte stem
cells.

Taken together, our observations on BrdU incorporation during regeneration,
on the position and number of dct+ cells during prolonged
MoTP incubation, and on the position and number of regenerated melanocytes in
kitj1e99 animals lead us to postulate that there are as
few as 15, or as many as 145, quiescent melanocyte precursors or stem cells in
the zebrafish larvae that contribute to the larval melanocyte regeneration
described here.

Our finding that the roles of the kit receptor tyrosine kinase in
the regeneration process are different from those previously described for the
ontogenetic development of larval melanocytes now raises the possibility that
the MoTP ablation and regeneration assay could be applied to forward genetic
analysis. This may allow us to specifically isolate mechanisms involved in the
regeneration process, including how the stem cell is kept in check by the
presence of its target tissue, or how the stem cell is activated to re-enter
the developmental pathway.

Acknowledgments

We thank Meera T. Sexena and John F. Rawls for critical reading of the
manuscript. We would also like to acknowledge Charles Higdon and Steven Jacob
for fish husbandry. The work was supported by National Institutes of Health
Grant R01-GM 56988 to S.L.J.

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